Abstract

The integrity of the genome is threatened by DNA damage that blocks the progression of replication forks. Little is known about the genomic locations of replication fork stalling, and its determinants and consequences in vivo. Here we show that bulky DNA damaging agents induce localized fork stalling at yeast replication origins, and that localized stalling is dependent on proximal origin activity and is modulated by the intra-S-phase checkpoint. Fork stalling preceded the formation of sister chromatid junctions required for bypassing DNA damage. Despite DNA adduct formation, localized fork stalling was abrogated at an origin inactivated by a point mutation and prominent stalling was not detected at naturally-inactive origins in the replicon. The intra-S-phase checkpoint contributed to the high-level of fork stalling at early origins, while checkpoint inactivation led to initiation, localized stalling and chromatid joining at a late origin. Our results indicate that replication forks initially encountering a bulky DNA adduct exhibit a dual nature of stalling: a checkpoint-independent arrest that triggers sister chromatid junction formation, as well as a checkpoint-enhanced arrest at early origins that accompanies the repression of late origin firing. We propose that the initial checkpoint-enhanced arrest reflects events that facilitate fork resolution at subsequent lesions.

Theoretical DNA structures and 2D–gel signals expected upon localized stalling of replication forks generated at a proximal origin or a distal origin. The replication forks stalled by DNA damage at an inactive origin site are generated at a distal origin (A). The stalling results in the formation of an early–Y arc, and a prominent fork accumulation signal at the stalling site. In contrast, the localized stalling of replication forks generated proximally at an active origin (B) results in the absence of an early–Y arc and the formation of a high rising bubble arc, along with a strong fork stalling signal. In both cases, once the DNA lesion is repaired or bypassed and replication resumes, nearly fully replicated fragments are expected to form a late–Y arc ending in a virtual 2N spot of structures with twice the original amount of DNA, while the unreplicated or fully replicated linear fragments migrate at the 1N spot.

Replication forks stall at early–firing origins in the presence of bulky DNA damage in S phase. G1–synchronized WT cells were released into S phase without DNA damaging agents (A) or in the presence of 1 μM adozelesin (B) or 0.15 μg/ml 4NQO (C). Samples were collected at the indicated time points post-release for 2D-gel electrophoresis and flow–cytometry analysis of DNA content. Replication intermediates were analyzed at three different regions encompassing early–firing origins ORI305, ORI508 and ORI1014 and were schematically represented in panel (D). The arrows in (B, C) indicate the major fork–stalling signals induced by adozelesin and 4NQO at the origins. Results for MMS–treated cells are shown in Supplementary Figure S1.

Replication fork stalling occurs in either direction at active origins. G1–synchronized WT cells were released into S phase in the presence of 0.15 μg/ml 4NQO. Samples were collected at 45 min post-release for 2D–gel electrophoresis. Replication intermediates were analyzed at ORI305 and ORI508 in the three alternative restriction digestion fragments indicated by the different colored bars. The relative position of the origin in each fragment is indicated by the diamond shape. The fork–stalling signal is indicated by arrows. Stalling–signal intermediates are schematically represented for each digest. Relevant signals observed in the presence of DNA damage are represented in the diagram. The bubble arc arising from an origin near the end of a DNA fragment is expected to be reduced in intensity and length and is schematized by a dotted line. Results for ORI607 and ORI306 are shown in Supplementary Figure S2.

Localized replication fork stalling requires proximal origin activity. (A) G1–synchronized ARS305-b5c7 mutant cells were released into S phase without DNA damaging agents or in the presence of 1 μM adozelesin or 0.15 μg/ml 4NQO. Samples were collected at 45 min post–release for 2D–gel analysis of replication intermediates at the inactivated ORI305 locus. (B) G1–synchronized WT cells were released into S phase without DNA damaging agents or in the presence of 1 μM adozelesin. Samples were collected at the indicated time points post-release for 2D-gel analysis of replication intermediates at two regions encompassing the naturally-inactive ARS304 and ARS302/303/320 situated adjacent to, and on the telomeric side of, ORI305 on chromosome III (map).

Heating of adozelesin-damaged DNA induces double-strand breaks at discrete sites. (A) G1–synchronized WT (panel 1–4) and ARS305–b5c7 mutant cells with inactive ORI305 (panel 5) were released into S phase in the presence of 1 μM adozelesin for 45 min and analyzed by 2D–gel electrophoresis. Replication intermediates were incubated in [10 mM Tris-HCl pH 7.9, 50 mM NaCl, 10 mM MgCl2, 0.1 mM EDTA] for 4 h at 65°C between the first and second dimension electrophoresis. The molecular size of the degradation DNA bands from the full-length restriction fragment (1N spot) and the position of the origin site within the fragment for ORI305 are annotated. A control sample prepared similarly from WT cells in an unperturbed S phase is shown for ORI305. Results for ORI508 are in Supplementary Figure S3. (B) The direction of replication forks within the ORI305 and ORI306 replicons and the relation between the presence of adozelesin adducts as observed in (A) and the localized stalling at active origins is schematically represented in WT cells (active ORI305, left) or in cells with inactivated ORI305 (right). Results for ARS304 and ARS302/303/320 loci in cells with inactivated ORI305 are shown in Supplementary Figure S3.

Inactivation of the intra–S–phase checkpoint diminishes damage–induced fork stalling at early–firing origins. G1–synchronized WT and mec1–100 cells were released into S phase in the presence of 1 μM adozelesin. Samples were taken for flow cytometry analysis (Supplementary Figure S4), and those collected at 30 and 45 min post–release for 2D–gel analysis of replication intermediates at early–firing origins ORI305 and ORI508 (WT, left panels for each ORI; mec1-100, right panels). The graphs show the fork–stalling signal intensity as a percentage of those of the replication intermediates in mec1–100 cells and separately in WT cells. Reduction in the relative fork stalling signal in the checkpoint mutant compared to that in WT cells was seen in three independent experiments, and also observed using 4NQO (Supplementary Figure S5), or using a ddc1Δ checkpoint mutant instead of mec1-100 (Supplementary Figure S6).

Inactivation of intra–S–phase checkpoint genes induces late origin firing, localized fork stalling and sister chromatid joining at ORI1412. (A) In control experiments, G1–synchronized WT cells were released into S phase without DNA damaging agents or in the presence of 1 μM adozelesin. Samples were collected at 45 min post–release for 2D–gel analysis of replication intermediates at late–firing origins ORI501 and ORI1412. (B) In checkpoint mutants mec1-100, ddc1Δ, tof1Δ and mrc1Δ, G1–synchronized cells were released into S phase in the presence of 1 μM adozelesin. Samples were collected at 45 min post–release for 2D–gel analysis of replication intermediates at late–firing origin ORI1412. The arrows indicate the X–shaped DNA structures and the fork–stalling signal induced in the checkpoint mutants.